P53/HDM2 and PERK/GCN2/Eif2α a Dissertation Presented to the Faculty of the C

The Roles of Two Different Pathways in Hypoxia: p53/HDM2 and PERK/GCN2/eIF2α

A dissertation presented to

the faculty of

the College of Arts and Sciences of Ohio University

In partial fulfillment

of the requirements for the degree

Doctor of Philosophy

Yan Liu

August 2009

This dissertation titled

The Roles of Two Different Pathways in Hypoxia: p53/HDM2 and PERK/GCN2/eIF2α

YAN LIU

has been approved for

the Department of Chemistry and Biochemistry

and the College of Arts and Sciences by

Susan C. Evans

Associate Professor of Chemistry and Biochemistry

Benjamin M. Ogles

Dean, College of Arts and Sciences 3

ABSTRACT

LIU, YAN, Ph.D., August 2009, Chemistry and Biochemistry

The Roles of Two Different Pathways in Hypoxia: p53/HDM2 and PERK/GCN2/eIF2α

(109 pp.)

Director of Dissertation: Susan C. Evans

Hypoxia always occurs in solid tumors. These hypoxic tumor cells are not sensitive to

chemotherapic agents because of poor drug delivery and slow proliferation. Hypoxia

activates two adaptive signaling pathways. On one hand, adaptation to hypoxia can be

regulated by hypoxia-inducible factor 1 (HIF-1) and its downstream genes. On the other

hand, hypoxia reduces protein synthesis and inhibits cell growth to adapt this stress. HIF-

1α plays a crucial role in tumor hypoxia and therapeutic resistance and its protein level is under tight control. Previous studies show that p53 suppresses HIF-1α protein through

HDM2-mediated ubiquitination and proteasomal degradation. However, the forced expression of HDM2 or growth factor-induced HDM2 can increase HIF-1α protein level, making it difficult to decipher how p53 and HDM2 regulate HIF-1α in hypoxia. In the first part of this study, we found that the increased p53 in hypoxia contributed to the downregulation of HIF-1α mRNA to suppress HIF-1α protein level. In addition, HIF-1α protein level was also inhibited by decreasing HDM2 protein level in hypoxia.

Furthermore, p53 and HDM2 knockout MEF cells were employed to determine the biological functions of p53 and HDM2 in hypoxia through modulating HIF-1α protein level. We showed that the presence of p53 inhibited hypoxia-induced cell growth arrest and cell cycle arrest through the suppression of HIF-1α and its downstream target, p21; 4

loss of HDM2 exhibited similar effects through the same mechanism; p53 strengthens

chemotherapeutic-induced apoptosis in hypoxia via downregulating HIF-1α, loss of

HDM2 displayed similar effects via the same mechanism. Recent studies suggest that

activation of PERK and phosphorylation of alpha subunit of eIF2 (eIF2α) confer cell

adaptation to hypoxic stress. However, eIF2α is still phosphorylated at a lowered level in

PERK knockout cells under hypoxic conditions. The mechanism for eIF2α kinase(s)-

increased cell survival is not clear. In the second part of this study, we investigated the

roles of GCN2-mediated eIF2α phosphorylation in hypoxia. Here we provided evidence

that another eIF2α kinase, GCN2, was also involved in hypoxia-induced eIF2α

phosphorylation. We demonstrated that both GCN2 and PERK mediated the adaptation to

hypoxic stress. High levels of eIF2α phosphorylation led to G1 arrest and protected cells

from hypoxia-induced apoptosis. Reduced phosphorylation of eIF2α by knocking out

either PERK or GCN2 suppressed hypoxia-induced G1 arrest and promoted apoptosis via

activation of p53 signal cascade. However, totally abolishing phosphorylation of eIF2α

inhibited G1 arrest without promoting apoptosis. In addition, reduced, but not abolished,

phosphorylation of eIF2α sensitized cells to chemotherapeutics, but not to gamma-

radiation in hypoxia. Based on our results, we propose that the levels of eIF2α phosphorylation serve as a “switch” in regulation of G1 arrest or apoptosis under hypoxic

conditions.

Approved: ______

Susan C. Evans

Associate Professor of Chemistry and Biochemistry 5

ACKNOWLEDGMENTS

I would like to express my gratitude to all those who assisted me in the research and who contributed to my knowledge from primary school to graduate school.

First I wish to thank my advisor, Dr. Susan Evans for her support throughout my research. She always encouraged me to think and work independently. This experience will help me during my whole academic career. Any future success will be owed to her patient guidance.

I would also like to thank my research committee members, Drs. Xiaozhuo Chen,

Jennifer Hines and Marcia Kieliszewski for their serving on my research committee. I will thank Dr. Shiyong Wu, our collaborator, for his exciting project and his guidance for the experiments and preparing the manuscript.

I enjoyed the environment in Dr. Evans’ lab and the friendship with my labmates:

Dr. Min Liang, Dr. Chrisanne Dias, Dr. Bernard Ayanga, Dr. Martin Schemerr, Eroica

Soans, Elroy Fernandes and Shuhua Du.

I would thank Department of Chemistry and Biochemistry and BMIT Program for their financial support. I would also like to thank Edison Biotechnology Institute for providing all the instruments.

I am grateful to Yanyan Cao, Yi Liu, Wei Liu and Dr. Csaba Laszlo for their contribution to my project and manuscripts.

Special thank to Dr. Xiaozhuo Chen and his wife, Dr. Yunsheng Li for their friendship making me not feel lonely. Finally, I thank my parents for their love and support. 6

TABLE OF CONTENTS

Page

Abstract ...... 3

Acknowledgments...... 5

List of Tables ...... 8

List of Figures ...... 9

Chapter 1: Introduction ...... 11

1.1 Tumor Hypoxia ...... 11

1.2 Hypoxia Inducible Factor 1 (HIF-1) ...... 13

1.3 Significance ...... 17

Chapter 2: Roles of p53 and HDM2 in Hypoxia by Modulating HIF-1 ...... 18

2.1 Introduction ...... 18

2.1.1 Tumor Suppressor p53 ...... 18

2.1.2 Oncogene Hdm2 ...... 23

2.1.3 Hypothesis ...... 26

2.2 Materials and Methods ...... 27

2.3 Results ...... 33

2.4 Discussion ...... 59

2.5 Future Directions ...... 66

Chapter 3: Roles of PERK and GCN2 in Hypoxia ...... 67

3.1 Introduction ...... 67

3.1.1 eIF2 and eIF2α phosphorylation ...... 67 7

3.1.2 Hypothesis and significance ...... 67

3.2 Materials and Methods ...... 69

3.3 Results ...... 71

3.4 Discussion ...... 85

References ...... 89

Appendix: List of abbreviations ...... 108

LIST OF TABLES

Page

Table 1: The transcriptional targets of HIF-1 ...... …. .16

Table 2: Estimated cancer cases with p53 mutations…………………………………...19 9

LIST OF FIGURES

Page

Figure 1: The roles of hypoxia in malignant progression in tumors…………………...... 12

Figure 2: Schematic representation of HIF-1α and HIF-1β………………………...... …14

Figure 3: The regulation of HIF-1α by oxygen levels…………………………………...15

Figure 4: The functional domains of p53………………………………………………...21

Figure 5: The regulation and function of p53……………………………………………22

Figure 6: The functional domains of HDM2………………………………………….....24

Figure 7: The effects of hypoxia on the protein levels of p53, HDM2 (or Mdm2 in MEF

cells) and HIF-1α……………………………………………………………………..34-35

Figure 8: The effects of hypoxia on HIF-1α at transcriptional and post-transcriptional

levels and its subcellular localization………………………………………………...36-37

Figure 9: Induction of p53 expression and transcriptional activity in hypoxia………38-40

Figure 10: Reduction of HDM2 at post-transcriptional levels in hypoxia…………...41-44

Figure 11: The effects of p53 and HDM2 (or Mdm2) on HIF-1α protein and mRNA levels………………………………………………………………………………….47-49

Figure 12: The effects of p53 and Mdm2 on cell proliferation and cell cycle distribution in MEF cells…………………………………………………………………………..52-54

Figure 13: Roles of p53 and Mdm2 in chemotherapeutics-induced apoptosis under hypoxic conditions……………………………………………………………………55-56

Figure 14: HIF-1 transactivation in normal cells and cancer cells………………………57

Figure 15: p53 transactivation in normal cells and cancer cells…………………………58 10

Figure 16: Models for the regulation of HIF-1α by p53 and HDM2 in hypoxia and how

p53 and HDM2 regulate cell proliferation, cell cycle distribution and apoptosis in

hypoxia by modulating HIF-1α protein level………………………………………...64-65

Figure 17: PERK and GCN2 affect cell survival in hypoxia………………………...73-75

WT PERK-/- GCN2-/- Figure 18: Hypoxia induces G1 arrest in MEF but not in MEF , MEF and

MEFA/A cells………………………………………………………………………….76-77

Figure 19: Knockout of PERK or GCN2 contributes to hypoxia-induced apoptosis……………………………………………………………………………...79-80

Figure 20: PERK and GCN2 are necessary for the recovery of cells from hypoxic stress……………………………………………………………………………………..82

Figure 21: The effects of cisplatin, actinomycin D, 5-fluorouracil and γradiation on cell

survival in both normoxia and hypoxia………………………………………………83-84 11

CHAPTER 1: INTRODUCTION

1.1 Tumor Hypoxia

Hypoxia is a common microenvironment in many human solid tumors as a result of the uncontrolled cell growth and the inadequate oxygen supply (Brown and Giaccia 1998;

Vaupel and Mayer 2005). Within tumor mass, about sixty percent of solid tumors show hypoxia (Vaupel and Mayer 2005), which contributes to tumor progression with time

(Vaupel and Mayer 2005) and resistance to both chemotherapy and radiotherapy (Brizel et al. 1999; Shasha 2001; Brown and Le 2002; Harrison et al. 2002). There are two major mechanisms underlying tumor propagation: genetic alterations and clonal selection

(Vaupel and Mayer 2005). Four genetic alterations have been observed, including point mutation, gene amplification, chromosomal aberration and polyploidy (Vaupel and

Mayer 2005). Mutations, such as either point mutations or deletion in p53, confer a growth advantage and mediate selection of resistant cells (Graeber et al. 1996). In addition to genetic alterations, hypoxia also mediates proteome changes resulting from posttranscriptional and posttranslational modification (Graeber et al. 1996). Therefore, tumor hypoxia may contribute to therapeutic resistance (Figure 1.1).

Hypoxia can activate signaling pathways mediating cell proliferation, angiogenesis and cell death. The in vitro studies demonstrate that cell proliferation is suppressed in hypoxia by reducing 40-50% of protein synthesis (Koumenis et al. 2002a) and/or inducing G1/S phase arrest (Giaccia 1996; Krtolica et al. 1999). Growth factors can be induced by hypoxia leading to new blood vessel formation (Harris 2002). In contrast, hypoxia can also induce apoptosis (programmed cell death) in p53 dependent or 12

independent manner. Severe hypoxia may trigger necrosis (necrotic cell death), which

can be observed in many human tumors (Soengas et al. 1999; Hockel and Vaupel 2001).

However, due to the changes in genome, tumor cells overcome oxygen deprivation and adapt to hypoxic conditions, leading to malignant progression with time (Vaupel and

Mayer 2005). In vitro studies demonstrate that oncogene transformed cells are more

resistant to hypoxia-induced cell death (Kim et al. 1997). In addition, hypoxia may also

mediate dedifferentiation of tumor cells (Axelson et al. 2005). Hence, the elements

involved in hypoxia signaling pathways may be good targets for cancer treatment.

Tumor Hypoxia and Malignant Progression

Malignant cell Viable cells Tumor hypoxia proliferation Diffusional O2

Angiogenic potential Genomic instability Apoptotic potential Clonal selection Resistance to therapy Proteome changes

Figure 1. The roles of hypoxia in malignant progression in tumors. Modified from Hockel and Vaupel, 2001 (Hockel and Vaupel 2001). 13

1.2 Hypoxia Inducible Factor 1 (HIF-1)

Hypoxia inducible factor 1 (HIF-1) is a key regulator in response to hypoxic stress. It

is a transcriptional factor, which consists of one alpha subunit and one beta subunit

(Wang et al. 1995; Wang and Semenza 1995). The transcriptional activity of HIF-1

depends on alpha subunit, which is regulated by oxygen levels and growth factors (Huang

et al. 1996; Agani and Semenza 1998; Laughner et al. 2001; Fukuda et al. 2002). HIF-1β, also known as aryl hydrocarbon nuclear receptor translocator (ARNT), is constitutively expressed (Wang et al. 1995). The human HIF-1α gene is located on chromosome 14

(14q21-q24), encoding an 826 amino acid protein. HIF-1β is on chromosome1 (1q21), encoding a 789 amino acid protein. Both alpha and beta subunits contain nuclear localization signals (NLS), a basic helix-loop-helix motif (bHLH), a Per-ARNT-Sim

(PAS) domain, and transactivation domains (TAD) (Figure 1.2). The bHLH and PAS domains mediate DNA binding and subunit dimerisation (Wang et al. 1995; Wang and

Semenza 1995). The TAD domains are responsible for HIF-1’s transcriptional activity

(Pugh et al. 1997). These domains also bind to some coactivators such as Ref-1 and p300/CBP (Wenger 2002). However, HIF-1α protein has a unique domain, oxygen- dependent-degradation domain (ODD), which is between 401 and 603 residues (Jiang et al. 1997; Huang et al. 1998). 14

OH OH Pro402 Pro564 HIF-1α

N- NLS bHLH PAS ODD N-TAD NLS C-TAD -C

DNA Binding Dimerisation Proteasomal Degradation

Nuclear Translocation Transactivation Nuclear Translocation

HIF-1β

N- NLS bHLH PAS TAD -C

Figure 2. Schematic representation of HIF-1α and HIF-1β. Modified from Bardos and Ashcroft, 2004 (Bardos and Ashcroft 2004).

When cells are in normoxia, proline 402 and proline 564 are hydroxylated by oxygen dependent proline hydroxylase. This modification recruits the binding of the von-

Hippel_Lindau protein (pVHL) and E3 ubiqutin ligase, which will degrade HIF-1α

(Figure 3) (Maxwell et al. 1999; Jaakkola et al. 2001). In hypoxia, the hydroxylation is inhibited and pVHL cannot bind to HIF-1α leading to its accumulation in hypoxic cells

(Jaakkola et al. 2001). Stabilized HIF-1α is translocated into the nucleus with HIF-1β, binds to hypoxic responsive elements (HREs), and activates downstream genes (Wenger

2002). In addition to oxygen, growth factors and oncogenes can also induce HIF-1α protein synthesis (Laughner et al. 2001). They include androgens, angiotensin II, EGF, herrgulin, HGF, IGF-1, insulin, interleujin-1β, thrombin, TNF- α and Ras (Bardos and

Ashcroft 2004). 15

pVHL Normoxia OH OH OH OH OH OH Pro402 Pro564 Pro402 Pro564 Pro402 Pro564 Pro402 Pro564

HIF-1α HIF-1α HIF-1α HIF-1α Ub Proline hydroxylase Ub Ub Ub

Hypoxia

Pro402 Pro564 Pro402 Pro564 Pro402 Pro564 Pro402 Pro564

HIF-1α HIF-1α HIF-1α HIF-1α Proline hydroxylase HIF-1β HIF-1β HRE

Figure 3. The regulation of HIF-1α by oxygen levels. Modified from Zarember and Malech, 2005 (Zarember and Malech 2005).

So far, more than sixty genes are found to be regulated by HIF-1, most of which facilitate angiogenesis, metabolic adaptation, cell cycle, proliferation, apoptosis and invasion/metastasis (Semenza 2003).

As HIF-1 is able to promote the production of growth factors and result in cell proliferation, it is considered to mediate tumor progression (Semenza 2003). In addition,

HIF-1α is overexpressed in tumors (Zhong et al. 1999; Zhou et al. 2006) and HIF-1 signaling pathways are involved in tumorigenesis (Carmeliet et al. 1998b). Knockdown of HIF-1α shows a reduced rate for tumor growth (Li et al. 2006a). Thus, it might be a potential target for tumor therapy (Lopez-Lazaro 2006).

Table 1

The transcriptional targets of HIF-1. Modified from Gaber et al., 2005 (Gaber et al. 2005).

Function Gene Angiogenesis and Erythropoiesis VEGF, VEGF receptor-1, TGFβ-3, EPO

Apoptosis and Cell Growth IGF-2, IGF binding protein-1, p21, Nix, Noxa, Nip3

Cell Migration and Matrix Metabolism SDF-1, c-Met, CXCR4, Collagen type V

Metabolism GLUT-1, GLUT-3, Aldolase A, C, LDH, HK-1

Iron Metabolism Transferrin, Transferrin receptor

Nucleotide Metabolism ETS1, DEC1, 2 pH control Carbonic anhydrase IX

1.3 Significance

Currently, two mechanisms have been discovered, through which tumor cells adapt to

hypoxia and survive in this microenvironment (Koumenis 2006a). The first one is

mediated by hypoxia inducible factor 1 (HIF-1), which can activate more than 60 genes

to induce angiogenesis and metabolism (Ratcliffe et al. 1998; Semenza 2000; Koumenis

2006a). Accordingly, HIF-1 is an essential regulator in tumorigenesis. In the first part of this study, we investigated how HIF-1 protein levels and transactivation are regulated by

p53 and HDM2 in hypoxia, as well as how p53 and HDM2 control cell proliferation, cell

cycle distribution, and apoptosis in hypoxia through modulating HIF-1 protein levels and

transactivation.

The second mechanism is mediated by inhibiting oxygen and energy consuming

process, including DNA replication and protein synthesis, which is in an HIF-1

independent manner (Hochachka et al. 1996; Sutherland et al. 1996; Hochachka and

Monge 2000; Koumenis 2006a). PERK and GCN2 phosphorylate eIF2α and thus reduce

mRNA translation initiation in response to stress (Koumenis and Wouters 2006a). In the

second part of this study, we determined how PERK and GCN2 affect cell adaptation to

hypoxia. 18

CHAPTER 2: ROLES OF P53 AND HDM2 IN HYPOXIA BY MODULATING HIF-1

2.1 Introduction

2.1.1 Tumor Suppressor p53

History

Thirty years ago, a 53kD protein (p53) was discovered, which binds to large-T

antigen of Simian Virus (SV-40) (Lane and Crawford 1979). Its encoding gene, p53, was

cloned from neoplastic human and rodent cells, which was then described to be an

oncogene because it had weak oncogenic properties (Harris 1996). Later in 1989, it was

realized that p53 cDNAs investigated by researchers were mutant forms rather than a

wild type form because they were cloned from mouse or human tumor cells. These

mutants contained missense mutations, causing p53 to act as a dominant-acting

oncogene. However, wild type p53 displays the ability to inhibit the transformation of

rodent fibroblasts in vivo, as well as the growth of human and mouse tumor cells in vitro

and in vivo (Harris 1996; Oren and Rotter 1999). Further studies discovered that p53 mutations occur in about 50% of various human cancers (Harris 1996). More than 20,000

mutations have been found in the p53 gene (Table 2). Rather than the commonly found

nonsense mutations, deletion, and insertions present in other tumor suppressor genes,

missense mutations are the dominant types in the p53 gene. The missense mutant p53

abrogates its tumor suppressive function and gains oncogenic function (Hussain and

Harris 2006).

Table 2

Estimated cancer cases with p53 mutations. Modified from Harris, 1996 (Harris 1996).

Cancer New cases Estimated cases with p53 mutations Breast 183400 44000

Colorectal 138000 68000

Lung 169900 95000

Lymphoma 24000 10400

Melanoma 34000 3000

Pancreatic 24000 10400

Prostate 244000 73000

Stomach 22800 9500

American Cancer Society, U.S. Estimates, 1995

Structure of p53

p53 encodes a 393 amino acid peptide (Fisher 2001), which contains several domains (Figure 4). p53 protein is a transcription factor (Polyak et al. 1997; Yu et al.

1999; Zhao et al. 2000; Fisher 2001; Yee and Vousden 2005). Its amino-terminal region is the transactivation domain, which interacts with the components of the transcriptional machinery such as TATA-binding protein (TBP), TBP-associated factors, and coactivator p300/CBP (Seto et al. 1992; Liu et al. 1993; Martin et al. 1993; Truant et al. 1993;

Horikoshi et al. 1995; Lu and Levine 1995; Thut et al. 1995; Avantaggiati et al. 1997; 20

Fisher 2001; Yee and Vousden 2005). This region is rich in proline, which is essential for

p53’s tumor suppressor function (Walker and Levine 1996; Venot et al. 1998). HDM2

also binds to this domain to negatively regulate p53 function by ubiquitinating and

degrading p53 (Haupt et al. 1997; Kubbutat et al. 1997; Kubbutat et al. 1998; Kubbutat

and Vousden 1998). However, the phosphorylation on some specific serine sites in this

domain, which is induced by various stress signals, such as ionizing radiation, can

alleviate the inhibition of p53 by HDM2 through the decreased interaction between these

two proteins (Shieh et al. 1997; Siliciano et al. 1997; Shieh et al. 1999). The central

domain of p53 contains the specific DNA binding sequence. Mutations within this

domain can disrupt the DNA binding activity of p53. Most human cancers result from

p53 mutants containing point mutations within this domain (Fisher 2001; Yee and

Vousden 2005). The carboxy-terminal domain contains nuclear export signal and nuclear localization signal, which are responsible for the cellular localization of p53 (Stommel et

al. 1999). This region also regulates the oligomerization of p53 (Sakamoto et al. 1994;

Jeffrey et al. 1995; Waterman et al. 1995), as well as obtains post-translational

modifications such as acetylation, phosphorylation, and sumolylation (Giaccia and

Kastan 1998; Yee and Vousden 2005).

1 97 300 393

N-terminal domain Central domain C-terminal domain Nuclear export signal Sequence-specific DNA binding Nuclear export signal Transactivation Nuclear localization signal Polyproline Oligomerization Phosphorylation sites Acetylation sites Mdm2 binding Phosphorylation sites Sumolylation sites

Figure 4. The functional domains of p53. Modified from Yee and Vousden, 2005 (Yee and Vousden 2005).

Function of p53

p53 knockout mice display very high tumor incidence and resistance to apoptosis

induced by ionizing radiation (Lowe et al. 1993; Lowe et al. 1994). In vitro studies show

that p53 is involved in various responses including cell growth, cell cycle arrest, DNA

repair, apoptosis and senescence. When cells are exposed to low stress, p53 induces G1/S checkpoint by activating its downstream genes, such as Cdk-inhibitor, p21(WAF1), and causes a transient growth arrest to repair the DNA damage by GADD45 and 14-3-3

(Alarcon-Vargas and Ronai 2002). However, when the DNA damage is severe, p53 induces genes, such as Bax, IGF-Bp3, Apo-1/Fas, PAG608 and KILLER/DR5 to trigger apoptosis and eliminate the risk of transformation (Miyashita et al. 1994; Buckbinder et al. 1995; Owen-Schaub et al. 1995; Israeli et al. 1997; Wu et al. 1997; Alarcon-Vargas and Ronai 2002). Overexpression of p53 also inhibits angiogenesis (Ravi et al. 2000). 22

Regulation of p53

p53 protein has a short half-life and is maintained at low levels by ubiquitin- mediated proteasomal degradation in normal cells under unstressed physiological conditions (Woods and Vousden 2001). HDM2 is a key negative regulator of p53 (Haupt et al. 1997). In response to various stresses, p53 is stabilized through post-translational modification, such as phosphorylation, and thus becomes activated. These stresses include DNA damage (UV and ionizing radiation, chemotherapeutics), hypoxia, cytokines, virus infection, heat shock and pH change (Kastan et al. 1991; Ashcroft and

Vousden 1999; Burns and El-Deiry 1999). Activated p53 may cause growth arrest, DNA repair or apoptosis (Figure 5).

p53-mediated responses

UV radiation

Ionizing radiation Growth arrest Hypoxia

DNA repair Cytokines p53 phospho-p53

Virus infection Apoptosis

Chemotherapeutics HDM2

Figure 5. The regulation and function of p53. Modified from Fisher, 2001 (Fisher 2001).

2.1.2 Oncogene Hdm2

Hdm2 (human homologue of mdm2) is classified as an oncogene because it is

overexpressed in various human tumors through three different mechanisms: gene

amplification (Oliner et al. 1992; Meddeb et al. 1996), enhanced transcription (Bueso-

Ramos et al. 1993; Watanabe et al. 1994), and increased translation (Landers et al. 1994;

Landers et al. 1997).

Structure of HDM2

Hdm2 gene encodes at least five polypeptides (p90-95, p85, p76, p74, p57-58)(Olson et al. 1993). The full length HDM2 protein consists of 491 amino acids and contains

several functional domains (Figure 6) (Marechal et al. 1997). The amino-terminal of

HDM2 contains the p53 protein binding site, nuclear localization signal (NLS), and nuclear export signal (NES) (Chen et al. 1993; Chen et al. 1995; Roth et al. 1998). Within the central domain is the acidic region, which binds to ribosomal protein L5 and 5S ribosomal RNA (Marechal et al. 1994). Within the carboxy-terminal domain is the conserved zinc finger region, which mediates protein-DNA and protein-protein

interaction. This region also possesses E3 ubiquitin ligase activity (Honda et al. 1997;

Honda and Yasuda 2000). RING finger domain also mediates protein-nucleic acid and

protein-protein interactions (Boddy et al. 1994; Elenbaas et al. 1996).

HDM2 Protein

p53 binding NLS NES Acidic domain Zinc finger RING finger

Figure 6. The functional domains of HDM2. Modified from Zhang and Wang, 2000 (Zhang and Wang 2000).

Function of HDM2

A primary function of HDM2 is to negatively regulate p53. Under unstressed

conditions, p53 does not increase the expression of HDM2 in normal cells (Mendrysa and

Perry 2000). HDM2 is required to maintain the low level of p53 and inhibit p53-induced apoptotic cell death (Marine et al. 2006). In HDM2 deficient mice with wild type p53, unrestricted p53 leads to early embryonic lethality, which can be rescued by additional deletion of p53 gene (Jones et al. 1995; Montes de Oca Luna et al. 1995). This

demonstrates that the inhibition of p53 by HDM2 is essential for embryonic

development. In contrast to unstressed conditions, p53 induces the expression of HDM2,

which in turn, binds, ubiquitinates, and degrades p53 in response to stress (Barak et al.

1993; Barak et al. 1994; Mendrysa and Perry 2000). In human cancers with wild type

p53, where HDM2 is amplified or overexpressed, such as in human soft tissue sarcomas

and breast cancer carcinoma, HDM2 inactivates tumor suppressor p53 (Cordon-Cardo et 25

al. 1994; Sheikh et al. 1993; Gudas et al. 1995; Marchetti et al. 1995; McCann et al.

1995).

Regulation of HDM2

The expression of HDM2 is regulated at three levels, including transcriptional, post-

transcriptional and post-translational regulation. Two distinct promoters have been

identified to regulate HDM2 transcripts. P1 promoter yields a longer mRNA transcript,

whereas the P2 produces a shorter mRNA transcript, which lacks 5’-untranslated regions

(Barak et al. 1994; de Oca Luna et al. 1996; Brown et al. 1999). P2 promoter contains two p53 binding elements, in which p53 binds and thus triggers the expression of HDM2

(Juven et al. 1993).

At least 40 different HDM2 mRNA transcripts have been discovered in normal cells and tumors (Bartel et al. 2002). Although some of their protein products are identified in tumors and possess the ability to promote the formation of tumors (Sigalas et al. 1996; Matsumoto et al. 1998; Evdokiou et al. 2001), most alternative spliced transcripts are unlikely translated into proteins due to the shift of HDM2 reading frame and thus are not correlated with prognosis (Oliner et al. 1992; Bartel et al. 2001). In contrast to the non-translated spliced mRNA products and oncogenic translated spliced protein products, when cells are treated with some various stress signals, such as UV and

IR, a certain HDM2 splice variant is induced to decrease the mRNA and protein levels of full length HDM2 and stabilize p53 (Chandler et al. 2006; Dias et al. 2006). Overall, the regulation of HDM2 at post-transcriptional level remains unclear. 26

In addition to the regulation at transcriptional and post-transcriptional levels,

HDM2, especially within amino-terminus, is likely to be phosphorylated by several

protein kinases (Mayo et al. 1997; Maya et al. 2001; Zhang and Prives 2001).

Phosphorylation on certain threonine residues of HDM2 can weaken the interaction between HDM2 and p53, thereby resulting in the decrease in p53 degradation by HDM2

(Zhang and Prives 2001). DNA damage-induced phosphorylation of HDM2 also accelerates its auto-ubiquitination and then auto-degradation (Stommel and Wahl 2004).

2.1.3 Hypothesis

Our study is based on the hypothesis that hypoxia modulates the levels of p53 and

HDM2 to regulate the level of HIF-1α protein, through which cell proliferation, cell cycle distribution, and apoptosis are controlled. Here, we present evidence that HIF-1α protein

level is affected by at least three different mechanisms that include p53, HDM2 and HIF-

1α protein itself: 1) Hypoxia-induced p53 suppresses HIF-1α mRNA expression; 2)

hypoxia-reduced HDM2 decreases HIF-1α protein level; 3) hypoxia-stabilized HIF-1α

protein inhibits its own mRNA expression. Through regulating HIF-1α level, p53 and

HDM2 modulate cell proliferation and cell cycle distribution in hypoxia. p53 and HDM2 also impact chemotherapeutic-induced apoptosis in hypoxia through the same mechanism. 27

2.2 Materials and Methods

Cell Culture Bronchial epithelial carcinoma cell line, H1299 (null for p53) was a gift from Dr. Jack Roth (M.D. Anderson Cancer Center, Houston, TX). H1299 p53 stable cell line (H1299 p53 OE) was established in our lab. Human breast epithelial adenocarcinoma, MCF-7 (wild-type p53) was purchased from ATCC (Rockville, MD).

MCF-7 p53 KD (p53 knockdown) was a gift from Dr. Ruiwen Zhang (University of

Alabama at Birmingham, Birmingham, Alabama). H1299 and MCF-7 cells were cultured in DMEM (Invitrogen, Carlsbad, CA) with 5% bovine growth serum (Hyclone, Logan,

UT) and 1% penicillin/streptomycin (Invitrogen, Carlsbad, CA). H1299 p53OE and

MCF-7 p53KD cells were cultured in the same medium described above with 400 μg/ml of G418 or 1 μg/ml of puromycine respectively. Human nontumorigenic immortalized cell lines, breast epithelial (MCF-12A), bronchial epithelial (NL20), were purchased from

ATCC (Rockville, MD). These two cell lines were cultured in DMEM/Ham’s F-12 (1:1) medium with 10% fetal bovine serum, 1% penicillin/streptomycin, 2.5 mM L-glutamine

(Hyclone), 10 mg/L insulin (Invitrogen), 0.001 mg/ml transferrin (Invitrogen), 20 ng/ml epidermal growth factor (Invitrogen), and 500 ng/ml hydrocortisone (Sigma, St. Louis,

MO).Mouse embryonic fibroblast (MEF) wild type (WT), mouse embryonic fibroblast p53-/- (1KO) and mouse embryonic fibroblast p53-/- mdm2-/- (2KO) were gifts from Dr.

Guillermina Lozano (University of Texas, M.D. Anderson Cancer Center, Houston, TX).

MEF cells were cultured in DMEM medium with 10% fetal bovine serum (Hyclone,

Logan, UT) and 1% penicillin/streptomycin. Hypoxic conditions were obtained using the

GasPak™ EZ Anaerobe Pouch System (BD Biosciences, VWR, S. Plainfield, NJ). 28

Oxygen levels were reduced to less than 1% (average 0.7%) in 2.5 hours. HIF-1α

expression plasmid, pcDNA3.1/V5-His-TOPO-HIF-1α, was a gift from Dr. Wei Gu

(Columbia University, New York, NY).

Plasmid purification The overnight cultures of E. coli were centrifuged at room temperature. The plasmids were purified using Qiagen Plasmid Purification Kit

(Qiangen). The pelleted bacterial cells were suspended in P1 buffer, P2 buffer and N3 buffer. The lysate was centrifuged and the supernatants were transferred to a spin column. The flow-through was discarded. The spin column was washed by PE buffer.

The plasmid DNA was eluted by EB buffer.

Transfection Transfection reagent lipofectamine 2000 (Invitrogen, Carlsbad, CA) was used for MCF-7 and H1299 cells and ExpressFectTM (Denville Sciemtific, Metuchen, NJ)

for MEF cells. 1×105 cells were plated in a 24-well plate with 500 µl of growth medium

24 hours before the transfection. Just prior to the transfection, the cells were washed

twice with DMEM without serum and antibiotics. 0.8 µg of plasmid was diluted in 50 µl

of DMEM and 2.0 µl of transfection reagent was diluted in 50 µl of DMEM. The diluted

plasmid and transfection reagent were incubated for 5 minutes at room temperature and

then were combined. The mixed solution was incubated for 20 minutes at room

temperature. One hundred microliters of the complexes were added to each well

containing DEME medium and cells. After 6 hours, full growth medium was added to the

o cells. The cells were incubated for 24 hours at 37 C in the CO2 incubator prior to further

analysis.

Western Blot Analysis Cells were lysed in buffer with 50 mM Tris-HCl pH 7.6, 150 mM 29

NaCl, 0.05% EDTA, 0.5% IGEPAL CA-630, and a cocktail of protease inhibitors (Roche,

Indianapolis, IN). Whole cell lysate was separated by SDS-PAGE gel and protein was transferred onto Immobilon-P membrane (Millipore). The membrane was blocked in 5% milk/PBS with 0.1% Tween-20. The blot was probed with the specific primary antibody and secondary antibody. Protein was detected by LumiGLO reagent and peroxide (Cell

Signaling, Danvers, MA). Anti-Mdm2 (N-20), anti-p53 (Bp53-12), anti-p21 (F-5) antibodies used were from Santa Cruz (Santa Cruz, CA). Anti-HIF-1α antibody was from

BD Biosciences (BD Biosciences Pharmingen, San Diego, CA). Anti-β-actin antibody was from Sigma (Sigma, St. Louis, MO). Anti-Akt and anti-phosphorylated Akt (Ser 473) were from Cell Signaling (Danvers, MA). Secondary horseradish peroxidase-linked antibodies were from Santa Cruz (Santa Cruz, CA).

Immunofluorescence Cells were seeded on slides prior to the treatment of hypoxia for 6 hours. After hypoxia treatment, cells were washed with cold PBS and fixed in ice cold

95% ethanol and 5% acetic acid at -20 oC for 5 minutes. Cells were incubated in PBS with 2% chicken serum for 30 minutes at room temperature and then were incubated in

PBS with 2% chicken serum and HIF-1α, p53 and HDM2 specific antibodies, H-206,

Bp53-12, SMP14 (Santa Cruz Biotechnology) for 1 hour. Then cells were incubated in

Texas red-conjugated secondary antibodies (Vector Laboratories, Burlingame, CA). The nucleus was stained by DAPI for 30 minutes. Photographs were taken using a Nikon microscope (West Chester, OH) and a digital camera (SPOT, DC Imaging) at 400 magnifications.

Cell Luciferase Assay Cells were seeded in a 24-well plate and transfected with HIF-1 30

luciferase reporter plasmid, pHIF-1-Luc (Panomics, Fremont, CA), Renilla luciferase reporter plasmid and other expression plasmids when applicable. The expression of luciferase was measured using the dual luciferase reporter assay kit (Promega, Madison,

WI). Cells were lysed and 20µl of the supernatant was analyzed. The intensity of the

luminescence was measured by a luminometer (Lumat LB 9507, Berthold, Oak Ridge,

TN). Renilla luciferase activity was used as a control to calculate relative activity.

RNA purification RNA was purified using RNeasy Mini Kit (Qiagen). Briefly, Cells

were washed by PBS twice and lysed by buffer RLT. The lysate was pipetted onto a

QIAshredder spin column. The same volume of 70% ethanol was added to the lysate. The

sample was pippeted onto a RNeasy mini column. The column was centrifuged and the

flow-through was discarded. The column was washed by buffer RW1. The column was

further washed by buffer RPE twice. RNA was eluted by RNase free water.

Real-Time Quantitative RT-PCR Assay Total RNA was extracted from cultured cells

using the RNeasy Mini Kit (Qiagen, Valencia, CA). First strand cDNA was obtained

using the iScript Select cDNA synthesis kit (Bio-Rad Laboratories, Hercules, CA). Equal

amounts of cDNA, appropriate primers and a master mix (iQ SYBR Green Supermix,

Bio-Rad Laboratories), were added together to make a 25 µl reaction mixture. The

thermocycler, iCycler iQ Real-Time Detection System (Bio-Rad Laboratories) was used

to perform PCR reactions. The β-actin mRNA was used as a control to compensate for

differences in RNA quantity between samples. hif-1 (mouse and human), p53, hdm2 and

β-actin (mouse) specific primers were purchased from SuperArray (Frederick, MD). β- 31

actin (human) specific primers were 5’-TGTGATGGTGGGAATGGGTCAG-3’ (sense) and 5’-TTTGATGTCACGCACGATTTCC-3’ (antisense).

Cell Proliferation Assay 2×103 cells were seeded in 96-well plates and then were exposed to normoxia or hypoxia. The cell number was determined by CyQUANT NF

Cell Proliferation Assay Kit (Molecular Probes, Invitrogen, Carlsbad, CA). The medium was removed from the cells. 50 μl of dye binding solution was dispensed into each well.

The plate was covered and incubated at 37 oC for 30 minutes. The fluorescence intensity

was measured by a microplate reader with the excitation wavelength at 485 nm and

emission wave length at 530 nm.

Cell Cycle Analysis Cells were cultured in serum free medium for 24 hours and were

trypsined. 106 cells were seeded in 10 cm plates and cultured in full medium for 24 hours.

The cells were exposed to normoxia or hypoxia for 24 hours. Treated cells were

harvested by trypsin and were washed by cold PBS. Cells were suspended in 200 μl of

PBS and fixed in 4 ml of cold 70% ethanol at -20oC overnight. Fixed cells were

centrifuged at 1000 rpm for 10 minutes and supernatant was aspirated carefully. Cells

were re-suspended in 100 µg/ml of RNase and 50 µg/ml of propidium iodide solution at

37oC for 30 minutes. 10,000 cells were analyzed by flow cytometry and cellquest software (Becton Dickinson, San Jose, CA). Cell cycle distributions were analyzed by

ModFit software (Verity Software House, Topsham, ME).

Apoptosis Assay 1×104 MEF WT, 1KO and 2KO cells were seeded in 96-well plates and

exposed to normoxia or normoxia plus 10 μg/ml of cisplatin or hypoxia plus 10μg/ml

cisplatin for 24 hours. The cells were analyzed using a Cell Death Detection ELISA kit 32

(Roche Diagnostics, Indianapolis, IN) to determine the cleaved DNA/Histone complex in

apoptotic cells. The assay was performed according to the manufacture protocol. Cells

were lysed and 20 μl of supernatant was transferred into a well. Immuno-reagent was

added into each well. The plate was incubated on a shaker at room temperature for 2

hours. Then substrate solution was pipetted into each well. The plate was incubated on a shaker for about 15 minutes. The absorbance was measured at 405/590 nm. 33

2.3 Results

Hypoxia significantly alters p53, HDM2 and HIF-1α protein levels in both transformed and nontransformed cells.

To investigate the effects of hypoxia on p53, HDM2 and HIF-1α, several cell lines were treated with hypoxia. In MCF-7 and H1299 p53 OE cells, where p53 is wild type or overexpressed, hypoxia induced an increase in p53 protein, a significant decrease in HDM2 protein, and a transient increase in HIF-1α protein (Figure 7A). In H1299 and

MCF-7 p53 KD cells, where p53 is null or knocked out, the effects of hypoxia on HDM2 and HIF-1α protein expression were similar to the cell lines with higher levels of p53

(Figure 7B). We also investigated wild type MEF cells (WT), and found that hypoxia showed similar effects on p53, Mdm2 (HDM2 murine homologue), and HIF-1α protein levels (Figure 7C). These results suggest that hypoxia showed similar effects on p53,

HDM2 and HIF-1α protein levels in both transformed and nontransformed cells. In addition, the expression profiles of HDM2 and HIF-1α were independent of p53.

We next examined the molecular mechanisms, by which hypoxia altered the protein contents of HIF-1α, p53, and HDM2. 34

A H1299 MCF-7 p53 KD

HIF-1α

HDM2

β-actin

Hypoxia 0h 6h 12h 24h 0h 6h 12h 24h

B MCF-7 H1299 p53 OE

HIF-1α

HDM2

p53

β-actin

Hypoxia 0h 6h 12h 24h 0h 6h 12h 24h

C MEF WT

HIF-1α

β-actin

MDM2

p53

β-actin

Hypoxia 0h 6h 12h 24h

Figure 7 The effects of hypoxia on the protein levels of p53, HDM2 (or Mdm2 in MEF cells) and HIF-1α. (A) p53 wild type MCF-7, p53 overexpression H1299 (H1299 p53 OE), (B) p53 null H1299, p53 knockdown MCF-7( MCF-7 p53 KD), and (C) p53 wild type MEF (WT) cells were exposed to hypoxia for the indicated times. Western blot analysis was carried out using p53, HDM2 and HIF-1α antibodies.

Hypoxia regulates HIF-1α protein contents via distinct mechanisms.

Hypoxia affects HIF-1α protein through post-translational hydroxylation

(Zarember and Malech 2005). However, as shown in Figure 7, hypoxia only induced a transient increase in HIF-1α protein. To address the question why a sharp decrease in

HIF-1α protein follows, we performed Real Time qPCR and surprisingly found that HIF-

1α mRNA decreased sharply during hypoxia in both H1299 and MCF-7 cells (Figure

8A). This leads to the question that in addition to the post-translational modification of

HIF-1α protein to stabilize itself, what factors also contribute to the transient increase in 36

its protein. As activated Akt (p-Ser 473) promotes HIF-1α protein synthesis, we analyzed

the phosphorylated Akt and total Akt in hypoxia. As shown in Figure 8B, phosphorylated

Akt levels increased, whereas total Akt levels kept constant. These results suggested that

hypoxia accumulated HIF-1α protein by promoting its protein synthesis by the induction of Akt phosphorylation. However, hypoxia could suppress HIF-1α mRNA expression. In addition, the stabilized HIF-1α protein translocated into the nucleus upon hypoxia (Figure

8C).

A 2.5 HIF-1α mRNA Expression in H1299 2

1.5

/actin α 1 HIF-1 0.5

0 Hypoxia 0h 6h 12h

1.5 HIF-1α mRNA Expression in MCF-7 1.2

0.9 /actin

α 0.6 HIF-1 0.3 0 Hypoxia 0h 6h 12h 37

H1299 MCF-7

p-Akt (Ser 473)

Akt

β-actin

Hypoxia 0h 6h 12h 24h 0h 6h 12h 24h

C HIF-1α

Normoxia

Hypoxia

Figure 8 The effects of hypoxia on HIF-1α at transcriptional and post-transcriptional levels and its subcellular localization. (A) H1299 and MCF-7 cells were treated with hypoxia for the indicated times. HIF-1α mRNA was analyzed by Real Time qPCR using specific primers. Data were from three independent experiments and standard deviation was determined. (B) Hypoxia induced Akt phosphorylation to trigger HIF-1α protein synthesis. Western blot analysis was carried out to show the levels of Akt serine 473 and 38

total Akt. (C) MCF-7 cells were exposed to either normoxia or hypoxia for 6 hours. After treatment, cells were probed by HIF-1α antibody and nucleus was stained by DAPI.

Hypoxia accumulates p53 protein by inducing its mRNA expression and phosphorylation on various serine residues.

A study reports that hypoxia stabilizes p53 protein through protein interaction between HIF-1α and p53 (An et al. 1998). To study whether hypoxia also impacts p53 mRNA expression, Real Time qPCR was performed in MCF-7 cells, where it was determined that an increase in p53 mRNA was observed after 6 hours of hypoxia treatment (Figure 9A). In addition, the phosphorylation on serine 9, 15, 20 and 37 of p53 was observed in hypoxia (Figure 9B). Notably, the phosphorylation on serine 6, 46 and

392 were not detectable in hypoxia. These results indicate that, in addition to the known stabilization of p53 by HIF-1α protein, hypoxia also induced p53 mRNA expression and phosphorylation on various serine residues. Then, hypoxia-induced p53 translocated into

the nucleus (Figure 9C) and exhibited an increase in transcriptional activity, which was

mediated by HIF-1α protein (Figure 9D). A 3.5 p53 mRNA Expression in MCF-7 3

2.5 2 1.5

p53/actin 1 0.5 0 Hypoxia 0h 6h 12h 24h 39

B MCF-7 p53

Phospho-Ser 9

Phospho-Ser 15

Phospho-Ser 20

Phospho-Ser 37

β-actin

Hypoxia 0h 6h 12h 24h

C p53

Normoxia

Hypoxia

D p53 Luciferase Activity in MCF-7 250 1400 1 200 1200 1 1000 1 150 800 pcDNA3 H RLU 100 RLU 600 400 1 50 200 0 0 0 Normoxia Hypoxia pcDNA3 HIF-1α

Figure 9 Induction of p53 expression and transcriptional activity in hypoxia. MCF-7 cells were treated with hypoxia for the indicated times. (A) p53 mRNA was analyzed by Real Time qPCR using specific primers. Data were from three independent experiments and standard deviation was determined. (B) Western blot analysis was carried out to show the phosphorylation levels of p53 on various serine residues. (C) MCF-7 cells were exposed to either normoxia or hypoxia for 6 hours. After treatment, cells were probed by p53 antibody and the nucleus was stained by DAPI. (D) The transcriptional activity of p53 was determined by p53 reporter plasmid. The p53 reporter plasmid was transfected into MCF-7 cells, followed by hypoxia treatment for 24 hours. The p53 transactivation was measured by the luciferase assay. Data were from at least three independent experiments and standard deviation was determined.

Hypoxia decreases HDM2 protein by accelerating the turnover of its protein.

To examine how hypoxia decreased HDM2 protein content, Real Time qPCR was performed which displayed that HDM2 mRNA levels did not change in hypoxia (Figure

10A). Previous studies suggest that HIF-1α protein contributes to the accumulation of p53 protein in hypoxia. To determine whether the decrease of HDM2 protein is due to the protein interaction between HIF-1α and HDM2, HIF-1α expression plasmid was transfected into H1299 and MCF-7 cells. The protein levels of HDM2 were determined 41

by western blot analysis, showing that HDM2 levels did not change (Figure 10B), which

indicates that the decrease of HDM2 protein was independent of HIF-1α protein. We next studied the half life of HDM2 protein in hypoxia. In contrast to HDM2 protein in normoxia, its half life was shortened in hypoxia in both H1299 and MCF-7 cells (Figure

10C). As HDM2 is degraded in the cytoplasm, the immunofluorescence assays displayed that some HDM2 protein was in the cytoplasm after 6 hours’ of hypoxia treatment compared to normoxia (Figure 10D). These results suggest that hypoxia accelerated the turnover of HDM2 protein, which was independent of p53 status.

A 1.4 HDM2 mRNA Expression in H1299 1.2 1 n 0.8

0.6 HDM2/acti 0.4 0.2 0 Hypoxia 0h 6h 12h 24h

1.6 HDM2 mRNA Expression in MCF-7 1.4 1.2 n 1 0.8 0.6 HDM2/acti 0.4 0.2 0 Hypoxia 0h 6h 12h 24h

B HIF-1α

H1299 MCF-7

HIF-1α

HDM2

β-actin

Normoxia HDM2 Hypoxia

1.2 H1299 1

Normoxia 0.8 Hypoxia 0.6

0.4 HDM2 remaining HDM2 0.2

0 0 5 10 20 30 40

CHX (Minutes) 44

1.2 MCF-7 1 Normoxia 0.8 Hypoxia 0.6

0.4 HDM2 remaining HDM2 0.2

0 0 5 10 20 30 40 CHX (Minutes)

Figure 10 Reduction of HDM2 at post-transcriptional levels in hypoxia. (A) H1299 and MCF-7 cells were treated with hypoxia for the indicated times. HDM2 mRNA was analyzed by Real Time qPCR using specific primers. Data were from three independent experiments and standard deviation was determined. (B) HIF-1α did not alter HDM2 protein levels. Different amounts of HIF-1α expression plasmids were transfected into H1299 and MCF-7 cells. Twenty four hours after transfection, HIF-1α and HDM2 were detected by western blot. (C) MCF-7 cells were exposed to either normoxia or hypoxia for 6 hours. After treatment, cells were stained using HDM2 antibody and DAPI. (D) H1299 and MCF-7 cells were exposed to hypoxia for 3 hours and then were treated with 100 μg/ml of cycloheximide (CHX) for the indicated times. Western blot was performed to determine HDM2 level.

p53 and HDM2 regulate HIF-1α protein levels via distinct mechanisms.

We then investigated whether the downregulation of HIF-1α protein and mRNA levels after 6 hours of hypoxia treatment is due to the increased p53 and the decreased

HDM2. p53 and HDM2 were increased by transfecting different amounts of their expression plasmids into H1299 and MCF-7 cells. Western blot results showed that the 45

increased p53 reduced HIF-1α protein contents while the increased HDM2 induced HIF-

1α protein contents (Figure 11A). To investigate whether endogenous p53 and HDM2

affect HIF-1α protein contents in the same way, we examined the HIF-1α protein in MEF

wild type (WT), MEF p53-/- (1KO) and MEF p53-/-mdm2-/- (2KO) (hdm2 murine

homologue) cells. Because mdm2 knockout is embryonically lethal, p53-/-mdm2-/- (2KO)

cells were compared to p53-/- (1KO) cells to study Mdm2’s function. Similarly, 1KO cells

were compared to WT cells to determine p53’s function. Our results showed that deletion

of p53 increased HIF-1α protein contents and further deletion of mdm2 decreased HIF-1α protein contents (Figure 11A).

To determine whether p53 and HDM2 (Mdm2) influence HIF-1α protein by modulating its mRNA expression, we transfected p53 or HDM2 expression plasmid into

MCF-7 cells and found that p53 repressed HIF-1α mRNA expression and HDM2 failed to impact HIF-1α mRNA expression (Figure 11B). We further employed MEF cells with different p53 and mdm2 status and observed that deletion of p53 induced HIF-1α mRNA expression while further deletion of mdm2 did not alter HIF-1α mRNA expression

(Figure 5B). These results demonstrated that p53 and HDM2 (Mdm2) altered HIF-1α

protein contents via distinct mechanisms. p53 decreased HIF-1α protein contents by the

suppression of HIF-1α mRNA expression, and HDM2 (Mdm2) increased HIF-1α protein

contents at post-transcriptional levels.

To study whether exogenous and endogenous p53 and HDM2 (Mdm2) alter HIF-1

transcriptional activity, we utilized p53 or HDM2 transfected H1299, MCF-7 and MEF

wild type cells, as well as MEF cells with different p53 and mdm2 status. The luciferase 46

assay results showed a decrease in HIF-1 transactivation by p53 and an increase in its

transactivation by HDM2 (Mdm2) in all the cell lines (Figure 11C). So far, our data

showed that hypoxia accumulated p53 protein via the upregulation of its mRNA

expression, decreased HDM2 protein via the acceleration of its own protein auto- degradation, and increased HIF-1α protein via the promotion of its protein synthesis.

Moreover, HIF-1α protein contents were tightly controlled by the alterations in p53 and

HDM2 levels in hypoxia via distinct mechanisms. 47

A HDM2 p53

H1299 MCF-7 H1299 MCF-7 HDM2 p53

HIF-1α HIF-1α

β-actin β-actin

MEF

HIF-1α

β-actin

WT 1KO 2KO

B HIF-1α mRNA in MCF-7 1.25 1 0.75 0.5 Fold Change Change Fold 0.25 0 pcDNA3 p53 HDM2

HIF-1α mRNA in MEFs 3 2.5 2 1.5 1 Fold Change Fold 0.5 0 WT 1KO 2KO

C 200 HIF-1 Transactivation 160

120 pcDNA3

RLU p53 80

0 H1299 MCF-7 50 HIF-1 Transactivation 40 pcDNA3 30 HDM2 HDM2 ALT1 RLU 20

0 H1299 MCF-7

0.8 HIF-1 Transactivation in MEF WT 0.6

0.4 RLU

0.2

0 pcDNA3 p53 HDM2

40 HIF-1 Transactivation in MEFs 30

20 RLU

0 WT 1KO 2KO

Figure 11 The effects of p53 and HDM2 (or Mdm2) on HIF-1α protein and mRNA levels. (A) Different amounts of p53 or HDM2 expression plasmids were transfected into H1299 and MCF-7 cells. Twenty-four hours after transfection, HIF-1α, p53 and HDM2 were detected by western blot analysis (left and middle). HIF-1α protein was detected by western blot analysis in MEF WT, 1KO, and 2KO cells after 24 hours of hypoxia treatment (right). (B) The p53 or HDM2 expression plasmid was transfected into MCF-7 cells. Twenty-four hours after transfection, the mRNA levels of HIF-1α were analyzed by Real Time qPCR (left). Data were from three independent experiments and standard deviation was determined. The mRNA level of HIF-1α was analyzed by Real Time qPCR (right) in WT, 1KO and 2KO cells cultured under normal conditions. Data were from three independent experiments and standard deviation was determined. (C) HIF-1 reporter plasmid was transfected into H1299, MCF-7, MEF WT, 1KO and 2KO cells with p53 or HDM2 expression plasmid if applicable. Twenty-four hours after transfection, HIF-1 transactivation was measured by the luciferase assay. Data were from at least three independent experiments and standard deviation was determined.

p53 and Mdm2 (HDM2 murine homologue) regulate proliferation, cell cycle distribution, and apoptosis in hypoxia by modulating HIF-1α levels.

Because the regulations of HIF-1α, p53 and HDM2 were similar in both human

cancer cells and MEF cells, MEFs with different p53 and mdm2 status were used as

models to assess the functional aspects of these three proteins/genes in hypoxia.

Hypoxia affects tumor growth; therefore, we performed a proliferation assay and found that cell growth rates were reduced in all cells during hypoxia compared to their normoxia cultured controls (Figure 12A), indicating that hypoxia resulted in cell growth

arrest in all cell lines. However, the most significant reduction in cell growth was

observed in p53-/- cells, followed by p53-/- mdm2-/- cells and then wild type cells. These

data suggest that p53 suppressed hypoxia-induced cell growth arrest and that Mdm2

promoted this arrest.

To explain this observation, cell cycle distribution was analyzed in MEF wild type

(WT), p53-/- (1KO) and p53-/- mdm2-/- (2KO) cells by exposing cells to hypoxia for 24 hours then measuring their DNA contents in each phase. In wild type (WT) cells cultured in hypoxia, G1 increased only 1.5% compared to the respective normoxic control;

-/- -/- -/- whereas G1 increased 19.2% in p53 (1KO) cells and 11.8% in p53 mdm2 (2KO) cells

compared to their respective normoxic controls (Figure 12B). These data suggested that

p53 inhibited hypoxia-induced G1 arrest and Mdm2 promoted this arrest, underlying the

results in Figure 12A.

We then further analyzed the cell cycle regulator HIF-1α and its downstream

target, p21. As expected, in hypoxia, the highest expression and inducibility of HIF-1α 51

protein compared to their respective normoxic controls were in p53-/- (1KO) cells,

followed by p53-/- mdm2-/- (2KO) and wild type (WT) cells (Figure 12C). Loss of p53 or

mdm2 led to a sharp decrease of p21 in normoxia (Figure 12C). However, hypoxia

treatment significantly induced p21 levels in these two cell lines. Among these three cell lines, p53-/- (1KO) cells showed the greatest inducibility of p21 levels in hypoxia

compared to those cultured in normoxia, followed by p53-/- mdm2-/- (2KO) cells and then

wild type cells, which was consistent with the cell cycle analysis results (Figure 12B).

Because p53 and Mdm2 (HDM2) do not regulate cell cycle in MEF HIF-1α-/- cells during

hypoxia (Goda et al. 2003), our data suggest that p53 and Mdm2 regulated hypoxia-

induced cell cycle arrest via HIF-1α and its downstream target, p21. p53 inhibited

hypoxia/HIF-1-induced G1 arrest through suppressing HIF-1α and p21 levels, and Mdm2

(HDM2) provoked this arrest through inducing HIF-1α and p21 levels. 52

Proliferation in 1 Day 1.4 1.2 1

0.8 Normoxia 0.6 Hypoxia

Fold Change 0.4 0.2 0 WT 1KO 2KO

Proliferation in 2 Days 1.4 1.2 1

0.8 Normoxia 0.6 Hypoxia

Fold Change Fold 0.4 0.2 0 WT 1KO 2KO

Proliferation in 3 Days 1.4 1.2 1

0.8 Normoxia 0.6 Hypoxia

Fold Change Fold 0.4 0.2 0 WT 1KO 2KO

Cell Cycle Distribution

80 78. 07 4. 73 17. 2 Normoxia77. 9Hypoxia 4. 19 17. 91 60 78. 23333 4. 286667 17. 48333 0. 438444 0. 403774 0. 376076 40 Percentage % Percentage 20

G1 S G2 G1 S G2 G1 S G2

WT 1KO 2KO

C MEF p21

β-actin Hypoxia 0h 6h 12h 24h

Hypoxia 24h - + - + - +

p21

β-actin

WT 1KO 2KO

Figure 12 The effects of p53 and Mdm2 on cell proliferation and cell cycle distribution in MEF cells. (A) MEF WT, 1KO and 2KO cells were exposed to hypoxia for different periods of time. After the treatment, cells were stained by DNA binding dye. The fluorescence intensity was measured by a microplate reader. Cells in normoxia were used as controls. Data were from three independent experiments and standard deviation was determined. (B) MEF WT, 1KO and 2KO cells were exposed to hypoxia for 24 hours. Cells were collected and stained with 50 µg/ml of propidium iodide. The cell cycle status was analyzed by a flow cytometer and ModFit software. Cells in normoxia were used as controls. Data were from three independent experiments and standard deviation was determined. (C) MEF WT, 1KO and 2KO cells were exposed to either normoxia or hypoxia for 24 hours. HIF-1α protein and p21 protein were analyzed by western blot.

Hypoxia/HIF-1α not only affects tumor growth, but also contributes to apoptosis.

Here, we investigated the roles of p53 and Mdm2 in chemotherapeutics-induced apoptosis during hypoxia via HIF-1α. Apoptosis was induced by cisplatin, ActD and 5-

FU in normoxia or hypoxia. As shown in Figure 13A, compared to their own normoxic controls, apoptosis in wild type (WT) and p53-/-mdm2-/- (2KO) cells increased 3-fold and 55

2-fold during hypoxia, respectively, whereas hypoxic p53-/- (1KO) cells had a 50%

decrease. Similar results were observed in ActD (Figure 13B) and 5-FU (Figure 13C) treated cells. Considering the expression and inducibility of HIF-1α protein in these cell

lines during hypoxia, the data suggest that p53 enhanced the effects of cisplatin, ActD

and 5-FU in hypoxia by decreasing HIF-1α protein levels. Furthermore, it suggests that

Mdm2 inhibited their effects by increasing HIF-1α protein levels.

A Cisplatin Induced Apoptosis 5 Normoxia 4 Hypoxia

Fold Change Fold 1

0 WT 1KO 2KO

B ActD Induced Apoptosis 3 2.5 Normoxia Hypoxia 2 1.5 1 Fold Change Fold 0.5 0 WT 1KO 2KO

C 5-FU Induced Apoptosis 2 Normoxia 1.5 Hypoxia

Fold Change Fold 0.5

0 WT 1KO 2KO

Figure 13 Roles of p53 and Mdm2 in chemotherapeutics-induced apoptosis under hypoxic conditions. MEF WT, 1KO and 2KO cells were treated with (A) 35 μM cisplatin, (B) 100 nM ActD (C) 100 μM 5-FU plus hypoxia. Cells treated with the same agent but cultured in normoxia were used as controls. After treatment, DNA/Histone complex was measured in apoptotic cells. Data were from three independent experiments and standard deviation was determined.

The balanced HIF-1 activity in cancer cells is disrupted by the deregulation of p53 and HDM2

To compare the activity of HIF-1 in normal cells and cancer cells, HIF-1 reporter plasmid was transfected into NL-20, H1299, MCF-12A and MCF-7 cells. HIF-1 luciferase assay showed that its activity was higher in H1299 and MCF-7 cells than in their nontransformed counterpart, NL-20 and MCF-12A cells, respectively (Figure 14).

NL-20 H1299 H1299 p53 0.822785 34.90789HIF-1 Transactivation 17.15493 0.792208 34.15662 18.83459 400.870968 33.98947 19.63359 NL-20 H1299 H1299 p53 age35 0.828654 34.35133 18.54104 v 0.039707 0.48919 1.265136 30

25NL-20 H12MCF-12A MCF-7 0.829 1.04 20 34.5 10.7 RLU 0.0397 0.0469 15 0.489 1.489 10

0 NL-20 H1299 MCF-12A MCF-7

Figure 14 HIF-1 transactivation in normal cells and cancer cells. HIF-1 reporter plasmid was transfected into NL-20, H1299, MCF-12A and MCF-7 cells. After 24 hours, cells were lysed then HIF-1 transactivation was analyzed by luciferase assay. Data were from at least three independent experiments and standard deviation was determined. To investigate the activity of p53 in wild type cancer cells and their nontransformed counterparts, p53 reporter plasmid was transfected into MCF-7 and MCF-12A cells. Although p53 is wild type in MCF-7 cells, the p53 luciferase assay showed that its activity was lower in MCF-7 cells compared to the nontransformed MCF- 12A cells (Figure 15).

13.9p53 101.54Transactivation MCF-7 MCF-12A 12013.88667 100.9567 1.390048 1.080386 100 80

60 RLU 40

0 MCF-7 MCF-12A

Figure 15 p53 transactivation in normal cells and cancer cells. p53 reporter plasmid was transfected into MCF-12A and MCF-7 cells. After 24 hours, cells were lysed and p53 transactivation was analyzed by luciferase assay. Data were from at least three independent experiments and standard deviation was determined.

2.4 Discussion

HIF-1 is a key player in hypoxic solid tumors (Wang et al. 1995). An understanding of how HIF-1 is controlled in hypoxia and how this process becomes deregulated in tumor cells are essential for basic research and clinical cancer research. Previous studies

have discovered that p53 and HDM2 regulate HIF-1 transactivation and its downstream

targets (Ravi et al. 2000; Bardos et al. 2004; LaRusch et al. 2007). HIF-1α protein binds

to p53 directly and is degraded by p53. p53 also mediates the binding of HDM2 to HIF-

1α, by which HDM2 mediates the ubiquitination and degradation of HIF-1α protein. This process is confirmed by the finding that overexpression of p53 increases the binding

between HIF-1α and HDM2 and thus augments the ubiquitination and proteasomal

degradation of HIF-1α protein (Ravi et al. 2000). However, later studies demonstrate that

it is HDM2 that binds to HIF-1α directly and that HDM2 acts as a bridge between p53

and HIF-1α (Chen et al. 2003; Nieminen et al. 2005). Forced expression of HDM2

enhances HIF-1 transactivation instead of ubiquitinating and degrading HIF-1α protein

(LaRusch et al. 2007). The degradation of HIF-1α by p53 is not mediated by HDM2.

Thus, an unanswered question is how p53 negatively regulates HIF-1. Moreover, all these

studies employed the overexpression of p53 and HDM2 in normoxia. For example, the

alterations of p53 and HDM2 level in hypoxia were not considered when addressing how

p53 and HDM2 affect HIF-1. In this study, we investigated the expression of HIF-1α, p53

and HDM2 in both transformed and nontransformed cells under hypoxic conditions, the

mechanism of how endogenous p53 and HDM2 regulate HIF-1α in hypoxia, and their

biological functions in hypoxia. 60

It is well established that hypoxia stabilizes HIF-1α protein and activates HIF-1 activity to promote angiogenesis and cell survival (Jiang et al. 1996; Li et al. 1996;

Salceda and Caro 1997). However, our in vitro studies showed a transient instead of a prolonged increase in HIF-1α protein levels during hypoxia (Figure 7A). Furthermore hypoxia did not induce HIF-1α mRNA expression. Instead, its mRNA showed a sharp downregulation (Figure 8A), which might explain the decrease in HIF-1α protein level after 6 hours of hypoxia. In addition to the stabilization of HIF-1α protein by hypoxia, the increase in phosphorylated Akt also contributed to the transient upregulation of HIF-1α protein in hypoxia (Figure 8B). These data suggest that hypoxia regulates the level of

HIF-1α by two different mechanisms. First hypoxia not only increases HIF-1α protein levels by inducing phosphorylated Akt to promote HIF-1α protein synthesis but also decreases HIF-1α mRNA to reduce HIF-1α protein. The second process is independent of p53 (Figure 8A). These regulations help in balancing the level of HIF-1α in hypoxia.

We next analyzed the level of p53 in hypoxia. Consistent with previous studies

(Alarcon et al. 1999; Hammond et al. 2002; Renton et al. 2003), our data showed that hypoxia induced p53 protein. The increased HIF-1α protein and HIF-1α-p53 interaction contribute to the induced p53 protein (An et al. 1998). Apart from this, the induced p53 mRNA and p53 phosphorylation on multiple serine residues also contributed to the increased p53 protein in hypoxia (Figure 9A and 9B). The phosphorylation of p53 at multiple serine sites in its C-terminus domain, where p53-HDM2 interaction occurs (Yee and Vousden 2005), alleviates the interaction between p53 with HDM2, thus inhibiting the degradation of p53 by HDM2 (Shieh et al. 1997). The translocation of p53 into the 61 nucleus and HDM2 into the cytoplasm in hypoxia confirmed that the phosphorylation of p53 disrupted their interaction (Figure 9C and 10C).

In response to stress, HDM2 is induced transcriptionally by p53 and in turn degrades p53 to balance its protein level and activity (Mendrysa and Perry 2000; Ringshausen et al. 2006). Although p53 was increased in hypoxia, HDM2 mRNA was not induced by p53 in response to hypoxic stress (Figure 10A). Additionally, HDM2 protein showed a significant decrease and was not detected after 24 hours of hypoxia in both transformed and nontransformed cells (Figure 7), suggesting that the regulation of HDM2 in hypoxia was not cell type specific. In contrast to various DNA-damage stresses, p53 could not transcriptionally induce HDM2 in response to hypoxic stress, because hypoxia induces p53 through a distinct mechanism other than from DNA-damage stress (Renton et al.

2003). HDM2 contains a nuclear export signal that helps in the shuttling of HDM2 to the cytoplasm, where it is degraded. In hypoxia, HDM2 shuttled to the cytoplasm (Figure

10C), which shortened the half life of HDM2 protein (Figure 10D). The accelerated auto- degradation of HDM2 may explain the decrease in HDM2 protein in hypoxia. In addition, unlike p53 protein (An et al. 1998), the stability of HDM2 protein in hypoxia was not relevant to HIF-1α protein (Figure 10B).

To study whether the decrease in HIF-1α protein and mRNA levels after 6 hours of hypoxia was due to the upregulation of p53 and the downregulation of HDM2, we analyzed the influence of p53 and HDM2 on HIF-1α protein and mRNA levels. Previous studies show that p53 binds to HIF-1α protein and degrades it through HDM2 (Ravi et al.

2000; Chen et al. 2003; Nieminen et al. 2005). However, in our study HDM2 was not 62

detected after 24 hours of hypoxia (Figure 7) suggesting that it could not mediate the interaction between p53 and HIF-1α. Here we proposed a novel mechanism, by which p53 and HDM2 regulate the level of HIF-1α. In hypoxia, p53 decreased HIF-1α protein level via the downregulation of HIF-1α mRNA level and HDM2 decreased HIF-1α protein level via the accelerated auto-degradation of HDM2 protein. Therefore, we can conclude that the alterations of p53 and HDM2 levels in hypoxia tightly regulate HIF-1α protein level via various mechanisms. Based on previous studies and our data, a model for the regulation of HIF-1α by p53 and HDM2 in hypoxia was proposed (Figure 16A).

Three factors modulate the levels of HIF-1α protein in hypoxia: 1) p53 increased to downregulate HIF-1α mRNA level; 2) HDM2 decreased to inhibit HIF-1α protein level;

3) HIF-1α protein suppressed its own level via the downregulation of its mRNA level.

Previous studies have described that p53 and HDM2 regulate angiogenesis through

HIF-1/VEGF (Ravi et al. 2000; Bardos et al. 2004; LaRusch et al. 2007). In addition to angiogenesis, the roles of p53 and HDM2 through HIF-1 in cell fate determination during hypoxia remain unknown. Therefore, we analyzed the growth characteristics and cell cycle distribution of MEF cells with different p53 and mdm2 status in hypoxia. HIF-1α is identified to be essential for hypoxia-induced cell cycle arrest and decreases cell growth rate by the activation of p21 in hypoxia (Carmeliet et al. 1998; Iida et al. 2002; Goda et al. 2003; Koshiji et al. 2004). Cells deficient of HIF-1α do not show any hypoxia-induced cell cycle arrest by p53 and HDM2 (Goda et al. 2003). Although p53 causes cell growth arrest and cell cycle arrest (Fisher 2001; Deb 2003), we found that p53 inhibited hypoxia/HIF-1-mediated cell growth arrest and cell cycle arrest by suppressing HIF-1α 63

(Figure 12A and 12B). In contrast, HDM2 could strengthen this arrest by the induction of

HIF-1α (Figure 10B). Therefore, our results provide a new insight into the functional consequences of the alterations of HIF-1α by p53 and HDM2 in hypoxia. In hypoxia,

HIF-1α accumulated, resulting in cell growth and cell cycle arrests. Meanwhile, p53 increased and HDM2 decreased, leading to the suppression of HIF-1α levels, the decrease in p21 levels, and the inhibition of hypoxia-induced cell growth arrest and cell cycle arrest (Figure 16B).

Cisplatin, ActD and 5-FU are common chemotherapy agents. Therefore, we explored the effects of p53 and HDM2 on chemotherapy agent-mediated apoptosis in hypoxia. A recent study reported that p53 contributes to chemo-resistance in ovarian carcinomas through cell cycle arrest and DNA repair (Moreno et al. 2007). However, in this study we found that loss of p53 led to a decrease in chemotherapeutics-induced apoptotic cell death in hypoxia; in contrast, loss of mdm2 augmented this cell death under the same conditions (Figure 13). The highest levels and the greatest inducibility of HIF-1α protein were observed in p53-/- cells, followed by p53-/-mdm2-/- cells and then wild type cells; the least sensitivity to the chemotherapeutics in hypoxia was also in p53-/- cells, followed by p53-/-mdm2-/- cells and then wild type cells. These results indicate that p53 and Mdm2 regulated chemotherapeutics-induced apoptosis in hypoxia, at least partially, by altering the levels of HIF-1α protein, because HIF-1α knockdown can increase the sensitivity to the therapies (Zhang et al. 2004; Zhang et al. 2004; Brown et al. 2006; Li et al. 2006;

Lopez-Lazaro 2006). 64

In summary, our results indicate that HIF-1α is under tight regulation by p53 and

HDM2 in both normal and cancer cells during hypoxia. These regulations influence cell growth, cell cycle distribution and apoptosis in hypoxia (Figure 16). However, these regulations are always damaged in human cancer cells. Mutation or loss of p53 occurs in more than 50% of human cancers (Freedman et al. 1999). Even in cancer cells with wild- type p53, p53’s activity was lowered (Figure 15), which might be due to the overexpression of HDM2 protein or the amplification of hdm2 gene (Freedman et al.

1999). These changes in HDM2 could also promote the activity of HIF-1 directly.

Accordingly, the activity of HIF-1 in cancer cells was higher than in their normal counterparts (Figure 14). The increased activity of HIF-1 through the downregulation of p53 and the upregulation of HDM2 can influence cell proliferation, cell cycle distribution and apoptosis. Our findings are significant to the elucidation of designing and utilizing therapeutic strategies for hypoxic solid tumors.

Hypoxia

HIF-1 mRNA p53 mRNA

HDM2 protein HIF-1 protein p53 protein

B Hypoxia

HDM2 p53

HIF-1 protein

p21

Hypoxia-Induced Chemotherapeutic–Induced Apoptosis Cell Cycle Arrest in Hypoxia

Figure 16 Models for the regulation of HIF-1α by p53 and HDM2 in hypoxia and how p53 and HDM2 regulate cell proliferation, cell cycle distribution and apoptosis in hypoxia by modulating HIF-1α protein level. (A) HIF-1α protein synthesis was promoted but mRNA expression was suppressed. p53 mRNA and protein increased and thus inhibited HIF-1α mRNA expression. HDM2 protein was decreased and inhibited HIF-1α protein. (B) p53 and HDM2 regulated cell cycle distribution through modulating HIF-1α and p21 levels. p53 and HDM2 also impacted the outcome of chemotherapeutics-induced apoptosis in hypoxia through the same mechanism. 66

2.5 Future Directions

We have demonstrated that hypoxia induced p53 to inhibit HIF-1 transactivation. In addition, hypoxia reduced HDM2 to suppress HIF-1 transactivation. Induced p53 and reduced HDM2 released hypoxia-induced G1 arrest and promoted apoptosis in hypoxia.

However, we have discussed that p53 is deactivated in many human cancers and HDM2 is overexpressed in some human cancers. Activating p53 and deactivating HDM2 might be a promising therapeutic for hypoxia solid cancers. To confirm our functional results in

MEF cells, human cancer cells with overexpressed p53 or knocked down HDM2 may be employed to determine the cell proliferation, cell cycle distribution, and apoptosis in hypoxia. 67

CHAPTER 3: ROLES OF PERK AND GCN2 IN HYPOXIA

3.1 Introduction

3.1.1 eIF2 and eIF2α phosphorylation

During the initiation of eukaryotic translation, at least nine eukaryotic initiation

factors (eIF) are involved in this process (Preiss and Hentze 1999; Sheikh and Fornace

1999; Koumenis 2006a). eIF2 is a heterotrimer composed of one alpha unit, one beta

subunit and one gamma subunit. GTP-bound eFI2 mediates the binding of tRNAMet to

40S ribosome. Once the initiation is finished, GTP is hydrolyzed into GDP and the initiation factors are released from the ribosome. GDP-bound eIF2 is exchanged for GTP- bound eIF2 with the help of the Guanine nucleotide exchange factor (GEF), eIF2B, to start another round of translation initiation. However, only unphosphorylated eIF2-GDP can be exchanged for GTP. At least four kinases can phosphorylate eIF2 at the alpha subunit on serine 51 induced by various stress, such as ER stress (RNA dependent protein kinase-like ER kinase, PERK), amino acid deprivation (general control non-depressible protein kinase 2, GCN2), hemoglobin deficiency (heme-regulated inhibitor kinase, HRI), and dsRNA presence (dsRNA induced protein kinase, PKR) (Kimball 1999; Dever 1999;

Kimball 1999; Kimball and Jefferson 2004; Hinnebusch 2005). Once phosphorylated, eIF2 is an inhibitor of eIF2B, which cannot exchange GDP for GTP, and the translation initiation is stopped (Duncan and Hershey 1989; Hershey 1989; Kaufman et al. 1989).

3.1.2 Hypothesis and significance

Recently, hypoxia has been reported to suppress protein synthesis and cell growth by activating PERK and phosphorylating eIF2α (Bi et al. 2005; Blais et al. 2006; Liu et 68

al. 2006; Koritzinsky et al. 2007). In vivo studies show that PERK promotes tumor adaptation and growth (Bi et al. 2005; Blais et al. 2006). These results suggest PERK and

eIF2α may be essential factors to regulate cell growth.

Four kinases, PKR, HRI, GCN2 and PERK, have been identified to phosphorylate

eIF2α and reduce translation initiation in response to stress (Koumenis and Wouters

2006b). Since both PERK and GCN2 can provide resistance to UV irradiation (Jiang and

Wek 2005), we tested the hypothesis that GCN2 might also play a role in response to

hypoxia. In contrast to wild type MEF cells, GCN2 knockout hampered cell hypoxia

adaptation in an eIF2α phosphorylation independent manner, like PERK knockout. This

is the first time that GCN2 was found to be involved in the hypoxia-induced

phosphorylation of eIF2α. Molecular mechanisms revealed that PERK or GCN2

mediated apoptosis independently of, but cooperatively with eIF2α phosphorylation. In

addition, PERK knockout, GCN2 knockout, or eIF2α mutation abolished hypoxia

induced cell cycle arrest, which was observed in wild type MEF cells, suggesting that

eIF2α phosphorylation was required for hypoxia induced cell cycle arrest. Moreover,

upon reoxygenation, PERK knockout, GCN2 knockout and eIF2α mutated MEF cells

exhibited lowered recovery from hypoxic stress, which indicated eIF2α phosphorylation

was essential to recovery. Finally, we observed that MEF cells lacking PERK or GCN2

were more sensitive to cisplatin in both normoxia and hypoxia. Taken together, these

findings established that both PERK and GCN2 were key regulators in response to

hypoxic stress. They might be good targets for anti-tumor therapies.

3.2 Materials and Methods

Cell Culture and Hypoxic Treatments MEF wild type (MEFWT), MEF PERK knockout

(MEFPERK-/-), MEF GCN2 knockout (MEFGCN2-/-), and MEF S51A mutant (MEFA/A) were kindly provided by Dr. RJ Kaufman (University of Michigan Medical School, Ann

Arbor, MI). They were cultured in DMEM (Cellgro, Herndon, VA) supplemented with

1% penicillin and streptomycin, 10% fetal bovine serum (Cellgro, Herndon, VA).

TM GasPak EZ Anaerobe Pouch System (BD Biosciences, VWR, S. Plainfield, NJ) was

utilized to obtain hypoxic conditions. Oxygen levels were reduced to 1% in 90 minutes.

Immunoblotting Treated cells were washed by cold phosphate buffer saline (PBS) twice

and lysed in buffer with 50 mM Tris-HCl, 150 mM NaCl, 0.05% EDTA, 0.5% IGEPAL

CA-630 and a cocktail of protease inhibitors. The same amount of total protein was

separated by SDS-PAGE gel. Protein was transferred onto Immobilon-P membrane. The

membrane was blocked in 5% milk in PBST (PBS with 0.1% Tween-20) at room temperature for 1 h. The member was probed with the specific primary and secondary antibodies in PBST. Protein was detected by LumiGLO reagent and peroxide (Cell

Signaling, Danvers, MA). The β-actin was used as a loading control. Anti-eIF2α, anti- phosphorylated eIF2α (Ser 51) and anti-β-actin were from Sigma (Sigma, St. Louis, MO).

Anti-Mdm2 (N-20), anti-p53 (Bp53), anti-HIF-1α and secondary horseradish peroxidase- linked antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA).

Cell Glucose Uptake Assay 3×104 MEF WT, 1KO and 2KO cells were washed with serum free DMEM twice. The cells were incubated in normoxia/hypoxia with serum free

medium for 24 hours. Cells were washed twice with KRP buffer (136 mM NaCl, 4.7 mM 70

KCl, 1.25 mM MgSO4, 1.25 mM CaCl2, 8.1 mM Na2HPO4, 1.9 mM NaH2PO4) and incubated with KRP buffer for 30 minutes. 50 µl of KRP supplemented with 1 µCi/ml

[3H] 2-deoxy-D-glucose and 1 mM glucose was added to each well and kept for 30

minutes. The buffer was discarded and the cells were washed with cold PBS. The cells were lysed with 0.2 M NaOH and transferred to a bottle. The radioactivity was measured

by LS 6500 multi-purpose Scintillation Counter (Beckman Coulter, Fullerton, CA). The

absolute value was quantified by the protein concentration.

Cell Viability Assay The cells (1x105) were seeded in 12-well plates and then were

exposed to normoxia or hypoxia. The viability was measured at different time points by

CellTiter-Glo reagent (Promega, Madison, WI). The medium was removed. 75 μl of water and 75 μl of CellTiter-Glo reagent were added into each well. The cells were incubated on a shaker for 5 minutes and at room temperature for 10 minutes. 100 μl of supernatant was measured by the luminometer (Lumat LB9507, Berthold Technologies).

Clonogenic Assay 5x103 cells were seeded in 6-well plates and cultured in 95% air and

5% CO2 at 37°C for 6 days. Cells were washed by PBS twice and fixed by cold methanol

at -20°C for 10 minutes. Fixed cells were stained by 1% crystal violet dissolved in 25%

methanol at room temperature for 10 minutes. The cells were then rinsed with distilled

water. The colonies, which have a size > 0.5 mm, were counted.

γ-irradiation The irradiation was performed by using a 137Cs irradiator (J.L. Shepherd

Associates).

3.3 Results

PERK and GCN2 protect cells from hypoxia-induced cell death independently of

eIF2α phosphorylation. Although the phosphorylation of eIF2α on Ser 51 in MEFPERK-/-

cells is reduced compared to MEFWT cells, the phosphorylation of eIF2α on Ser 51 still occurs (Liu et al. 2008), which indicates the involvement of other EIF2AK part from

PERK. To determine whether GCN2 is also involved in hypoxia-induced phosphorylation of eIF2α, a GCN2 knockout cell line, MEFGCN2-/-, was used to compare

PERK-/- WT to MEF and MEF cells. The time-dependent phosphorylation of eIF2α in the four

cell lines was determined after hypoxia-treatment. In contrast to the MEFWT cells, the

hypoxia-induced phosphorylation of eIF2α was delayed in MEFPERK-/- cells (Figure 17A,

partly performed by Dr. Csaba Laszlo), which agreed with previous reports (Koumenis et

al. 2002b; Liu et al. 2008). As expected, there was also a delay of the phosphorylation of

eIF2α in MEFGCN2-/- cells (Figure 17A). This is the first time that GCN2 was found to be

involved in the hypoxia-induced phosphorylation of eIF2α. In addition, this is the first

demonstration that GCN2 is also involved in the hypoxia-induced phosphorylation of

eIF2α.

Hypoxia-induced PERK activation protects cells from death by increasing eIF2α

phosphorylation (Bi et al. 2005). To determine whether GCN2 can also protect cells

from hypoxia-induced cell death, we analyzed viabilities of MEFWT, MEFPERK-/-,

MEFGCN2-/- and MEFA/A cells under normoxic or hypoxic conditions. Our data showed

that under normoxic conditions, the growth rates of MEFWT, MEFPERK-/-, MEFGCN2-/- and

MEFA/A were within 10% differences (Figure 17B, Normoxia), but under hypoxic 72

conditions, viabilities of MEFGCN2-/- and MEFPERK-/- cells decreased more rapidly than

MEFWT and MEFA/A cells (Figure 17B, Hypoxia). After correcting the survival rates in

hypoxia with the growth rates in normoxia, our data demonstrated that the survival rates for MEFGCN2-/- and MEFPERK-/- cells were decreased to 2.4±0.1% and 5.1±0.3%,

respectively (Figure 17B, Hypoxia/Normoxia). Surprisingly, the survival rate of MEFA/A cells was 47.3±3.2%, which was similar to the 44.9±2.1% survival rate of MEFWT cells

(Figure 17B, Hypoxia/Normoxia). These results suggest that PERK and GCN2 protect cells from hypoxia-induced death independently of eIF2α phosphorylation levels.

To further confirm our conclusion, we analyzed cell viabilities using metabolic and morphologic analyses. Glucose uptake assay was used to measure the metabolic activities of the cells in hypoxia. Our data showed that while the metabolic activities of MEFWT and MEFA/A cells were maintained at 75.4±0.3% and 83.5±0.4%, respectively, the

activities in MEFGCN2-/- and MEFPERK-/- cells were decreased to 36.0±0.2% and

42.3±0.2%, respectively (Figure 17C, performed by Yi Liu). In addition, the

morphological analysis also showed that the MEFGCN2-/- and MEFPERK-/- cells were more

sensitive to hypoxia than MEFWT and MEFA/A cells (Figure 17D). These data support our

conclusion that both PERK and GCN2 protect cells from hypoxia-induced cell death in

an eIF2α phosphorylation independent manner.

Figure 17 PERK and GCN2 affect cell survival in hypoxia. (A) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to hypoxia for the indicated time points before immunoblotting with phosphorylated eIF2α (Ser 51) and total eIF2α specific antibodies. (B) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to normoxia or hypoxia for indicated time points and then viability assays were performed. 75

The bars represent the means of three independent experiments. (C) Hypoxia lowers glucose uptake much more in MEFPERK-/- and MEFGCN2-/- cells. MEFWT, MEFPERK-/- and MEFGCN2-/- and MEFA/A cells were exposed to hypoxia for 12 hours with serum free DMEM. Glucose supplemented with [3H] 2-deoxy-D-glucose was added. Total protein was collected and the radioactivity was measured by liquid scintillation spectrometry. The experiments were performed in triplicate. (D) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to normoxia or hypoxia for indicated time points. Cells were photographed using microscopy and supplied Nikon digital camera.

Hypoxia-induced cell cycle arrest is dependent on PERK and GCN2. To determine whether both PERK and GCN2 play a role in regulation of cell cycle arrest in hypoxia, we performed cell cycle analysis in MEFWT, MEFGCN2-/-, MEFPERK-/- and MEFA/A cells.

WT PERK-/- Our data showed that hypoxia induced G1 arrest in MEF cells but not in MEF ,

MEFGCN2-/- or MEFA/A cells (Figure 18A and 18B). These data demonstrate that the

activation of PERK/GCN2 and the phosphorylation of eIF2α were essential for hypoxia- induced cell cycle arrest.

To access the molecular mechanism for PERK/GCN2-mediated eIF2α phosphorylation in regulation of cell cycle arrest, we analyzed the expressions of key cell cycle regulators, HIF-1α and p21 in those cells. Our data showed that HIF-1α expression was induced most significantly in MEFWT cells compared to MEFPERK-/- and MEFGCN2-/- cells, and it was not detected in MEFA/A cells (Figure 18C). In contrast to HIF-1α, the

maximal inducibility of p21 in hypoxia was achieved in MEFWT cells (Figure 18C). The

inducibility, but not the expression levels of p21, was correlated to the hypoxia-induced

cell cycle arrest. These results suggest that the hypoxia-induced cell cycle arrest is

mediated by a transient induction of p21, which is mediated by PERK/GCN2. 76

A Normoxia Hypoxia

WT WT

PERK-/- PERK-/-

-/- GCN2-/- GCN2

A/A A/A

WT PERK-/- GCN2-/- Figure 18 Hypoxia induces G1 arrest in MEF but not in MEF , MEF and MEFA/A cells. (A-B) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were treated with hypoxia for 12 hours before the performance of cell cycle analysis using PI staining and FACS machine and Cellquest software. Results were analyzed by ModFit software. Cells cultured in normoxia were also analyzed as controls. The bars represent the means of three independent experiments. (C) Cell cycle regulators were analyzed by western blot. MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were treated with hypoxia for the time points as indicated and immunoblotted with anti-HIF-1α and anti-p21 specific antibody.

PERK and GCN2 protect cells from hypoxia-induced apoptosis. To determine

whether GCN2, like PERK, protects cells from hypoxia-induced apoptosis, we accessed

the roles of PERK/GCN2-mediated eIF2α phosphorylation in regulation of apoptosis in

hypoxia. MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to hypoxic stress and then apoptotic cell death was analyzed by examining the cleavage of histone-

DNA complex. Our data showed that much higher levels of nucleosome contents were displayed in MEFPERK-/- and MEFGCN2-/- cells than the levels in MEFWT and MEFA/A cells

(Figure 19A).

p53 plays a central role in regulation of apoptosis during hypoxia (Fisher 2001).

Therefore, we determined whether PERK and GCN2 would protect cells from hypoxia- induced apoptosis via regulating p53 signaling pathway. Our data demonstrated that p53 transcriptional activities were increased 4.0±0.5 and 4.4±0.8 folds in MEFPERK-/- and

MEFGCN2-/- cells, respectively, while the activities were only increased 1.4±0.1 and

1.7±0.1 folds in MEFWT and MEFA/A cells, respectively, in hypoxia (Figure 19B). We

also examined the protein level of p53 by western blot, which showed that p53 was upregulated most significantly in MEFPERK-/- and MEFGCN2-/- cells (Figure 19C). The

transcriptional activities of a p53 downstream gene, Bax, were also up regulated more in

MEFGCN2-/- and MEFPERK-/- cells than in MEFWT and MEFA/A cells (Figure 19D). At the

same time, the expression of a p53 negative regulator, Mdm2, was decreased in all four

cell lines independent of eIF2α phosphorylation levels (Figure 19E). These results

suggest that activation of PERK and GCN2 inhibits hypoxia-induced activation of p53

signaling pathway independent of Mdm2. 79

A Apoptosis

B p53 Transactivation

D Bax Promoter

Figure 19 Knockout of PERK or GCN2 contributes to hypoxia-induced apoptosis. (A) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to normoxia or hypoxia for 36 hours before performing ELISA cell death assays by detecting cleaved Histone/DNA complex in apoptotic cells. The bars represent the means of three independent experiments. Apoptotic regulators were analyzed by luciferase assay (B-C) and western blot (D). MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were cotransfected with p53 luciferase reporter plasmid (B) or Bax luciferase reporter plasmid (C). Renilla luciferase reporter plasmid was also used to normalize the transfection efficiency. After 24 hours, cells were exposed to normoxia or hypoxia for 12 hours before the performance of luciferase assay. The bars represent the means of three independent experiments. MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were treated with hypoxia for the time points as indicated and immunoblotted with anti-Mdm2 antibody (D). 81

PERK and GCN2 promote the recovery of cells from hypoxia. The ability to recover from stress is also an important property to measure the regulation of survival and growth of cells after exposure to stress. We determined whether PERK/GCN2-mediated eIF2α phosphorylation affects the ability of cells to recover from hypoxia. Clonogenic assay was used to measure survival and recovery of cells from hypoxic stress. The MEFWT,

MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to hypoxia for 24 hours and then

cultured under normal conditions for 6 days (Figure 20A). Our data showed that,

compared to MEFWT cells, the recovery rates after hypoxia were reduced 50.5±8.4% and

50.0±2.9% in MEFPERK-/- and MEFGCN2-/- cells, respectively (Figure 20B). The recovery

rate of MEFA/A cells was only reduced 5.9±5.9% (Figure 20B), which was consistent with our viability results (Figure 17B). Noticeably, the sizes of MEFA/A colonies were much

smaller for the other three cell lines. The results suggest that the hypoxia-induced

activation of PERK and GCN2 promotes cell recovery from hypoxia. However, recovery

rates after hypoxia are not totally correlated with the levels of eIF2α phosphorylation.

Complete deletion of eIF2α phosphorylation did not affect recovery rate but influence the

cell growth rate after hypoxic stress. 82

Figure 20 PERK and GCN2 are necessary for the recovery of cells from hypoxic stress. (A) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were exposed to normoxia or hypoxia for 24 hours. Cells were collected and stained by Trypan Blue. Non-apoptotic cells were counted. Five thousand cells were re-plated and cultured under normoxia for 6 days. Cells were photographed using microscopy and supplied Nikon digital camera. (B) Percentage of MEFWT was determined. The bars represent the means of three independent experiments.

Knocking out PERK or GCN2 sensitizes MEF cells to chemotherapy but not radiotherapy in normoxia and hypoxia. p53 plays important roles in mediating 83 cytotoxicity induced by chemotherapeutics (cisplatin, 5-FU and ActD) and γ-radiation. p53 was upregulated much more in MEFPERK-/- and MEFGCN2-/- cells during hypoxia.

Therefore, we determined whether these two cell lines would be more sensitive to chemotherapeutics in hypoxia. Our data showed a sharp decrease of viability in MEFPERK-

/- and MEFGCN2-/- cells compared to MEFWT and MEFA/A cells under both normoxic and hypoxic conditions (Figure 21A-C). In contrast, all four cell lines responded to γ- irradiation in a similar manner during normoxia and hypoxia (Figure 21D). A

Figure 21 The effects of cisplatin, actinomycin D, 5-fluorouracil and γradiation on cell survival in both normoxia and hypoxia. MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were cultured treated with (A) 15 μM cisplatin, (B) 50 μM 5-FU, or (C) 30 nM Act D and then cultured under normoxia or hypoxia for 24 hours before performing viability assay. The bars represent the means of three independent experiments. (D) MEFWT, MEFPERK-/-, MEFGCN2-/- and MEFA/A cells were treated with 10 Gy γ-radiation and exposed to normoxia or hypoxia for 24 hours before performing viability assay. Cells cultured in normal medium without γ-radiation treatment were used as controls. The bars represent the means of three independent experiments. 85

3.4 Discussion

One response to hypoxia is to decrease macromolecular synthesis and slow down cell proliferation (Koumenis 2006; Koumenis and Wouters 2006b). Recent studies show that hypoxia can activate PERK. Then PERK phosphorylates eIF2α, reduces protein

synthesis, and contributes to hypoxia adaptation (Koumenis et al. 2002; Bi et al. 2005;

Blais et al. 2006; Liu et al. 2006; Koritzinsky et al. 2007; Liu et al. 2008). However, in

hypoxic cells lacking PERK, the phosphorylation of eIF2α on Ser 51 is still upregulated

(Liu et al. 2008), which indicates that there is another kinase that can phosphorylate

eIF2α on Ser 51 in hypoxia. Previous studies report that both PERK and GCN2 can

phosphorylate eIF2α in response to UV (Marbach et al. 2001; Deng et al. 2002; Jiang and

Wek 2005). GCN2 is an eIF2α kinase and it is controlled by amino acid abundance

(Berlanga et al. 1999; Sood et al. 2000). GCN2 can also upregulate the mRNA translation

of GCN4, which positively regulates amino acid synthesis (Ramirez et al. 1991; Qiu et al.

1998; Wek et al. 2006). Here, we found, upon hypoxia, the phosphorylation of eIF2α in

MEFGCN2-/- cells was also delayed, like MEFPERK-/- cells, suggesting GCN2 is also

involved in the phosphorylation of eIF2α in response to hypoxia. This is the first time

that GCN2 was reported to be involved in hypoxia.

To study whether both PERK and GCN2 are involved in adaptation to hypoxic

stress, we exposed MEFPERK-/-, MEFGCN2-/- and MEFWT cells to normoxia and hypoxia and

compared their survival rates. Interestingly, MEFPERK-/- and MEFGCN2-/- displayed almost

the same survival rates, which were much lower than MEFWT cells. These results

indicated that PERK and GCN2 play the same roles in response to hypoxia. Previous 86 studies show that eIF2α mutation can also reduce cell survival in hypoxia (Bi et al. 2005).

We subsequently analyzed whether PERK and GCN2 regulate cell adaptation via eIF2α phosphorylation. Our results demonstrated that eIF2α phosphorylation was not required.

To confirm these data, we further determined the biological effects of PERK, GCN2 and the phosphorylation of eIF2α on glucose uptake in response to hypoxia. We found that knockout of PERK or GCN2 impaired glucose uptake in hypoxia independent of eIF2α phosphorylation level. These data partially proved our hypothesis that the action of

PERK and GCN2 is independent of the phosphorylation level of eIF2α on Serine 51.

Although previous in vivo studies show tumors derived from mutated eIF2α MEF cells are smaller than tumors from wild type MEF cells, tumors from PERK knockout MEF cells are extremely smaller than tumors from mutated eIF2α MEF cells (Bi et al. 2005).

In addition, knockout of PERK can downregulate angiogenesis (Blais et al. 2006), but eIF2α mutation has no effects on angiogenesis (Bi et al. 2005). Therefore, previous studies suggest that PERK regulate cell adaptation in an eIF2α independent manner. Our results demonstrated GCN2 promotes cell survival independent of eIF2α phosphorylation level.

Hypoxia affects cell survival via growth arrest and apoptosis via HIF-1α (Goda et al. 2003b; Koshiji et al. 2004). In this study, we analyzed whether PERK and GCN2 regulate cell survival via cell cycle arrest and apoptosis. In cells lacking PERK or GCN2, hypoxia-induced cell cycle arrest was hampered compared to wild type cells. Although the total p21 protein in MEFPERK-/-, MEFGCN2-/- , MEFA/A and MEFWT cells was upregulated, the induction of p21 during hypoxia in MEFWT cells was most significant, 87

which could explain the cell cycle arrest in MEFWT cells. Therefore, our data suggest that

another pathway may cause cell cycle arrest via p21 in hypoxia. Another group has

reported that after protein response pathway (UPR) is activated, PERK and GCN2

cooperatively regulate the phosphorylation of eIF2α and the subsequent G1 phase arrest

(Hamanaka et al. 2005). Although we used different stress, our results support that the

phosphorylation of eIF2α is essential for hypoxia or other stress induced cell cycle arrest.

Our further studies showed that, like PERK, knockout of GCN2 also promoted

hypoxia induced apoptosis. To the best of our knowledge, this is the first time that

knockout of GCN2 induces apoptosis in response to hypoxia. Since we have confirmed

that PERK and GCN2 regulated cell cycle via the phosphorylation of eIF2α, we also

tested whether the apoptosis would be via the phosphorylation of eIF2α. We found that

MEFA/A and MEFWT cells exhibited the same levels of hypoxia induced apoptosis at

different time points. Other groups also find PERK and GCN2 can affect cell adaptation independently of eIF2α. In response to UV or glucose stimulation, GCN2 has no effects

on the upregulation of phosphorylation of eIF2α or the translation of GCN4 mRNA

(Marbach et al. 2001). A novel function of PERK has been identified to mediate UV

induced and force induced apoptosis in an eIF2α independent manner (Parker et al. 2006;

Mak et al. 2008). In the case of hypoxia, PERK and GCN2 increase cell tolerance via a

novel mechanism.

In addition to the roles of PERK and GCN2 in hypoxia induced cell cycle arrest and

apoptosis, their effects on the cell recovery from hypoxia were also determined. Upon

reoxygenation, MEFPERK-/- and MEFGCN2-/- cells showed the most reduced recovery, 88

whereas MEFWT cells demonstrated full recovery. In addition, the recovery in MEFA/A

cells was apparent but less significant than that in MEFPERK-/- and MEFGCN2-/- cells. The

best recovery in MEFWT cells may be due to the cell cycle arrest. These results suggest

that cell cycle arrest protects cells from being impaired. The eIF2α independent functions

of PERK and GCN2 can explain the slower recovery in MEFPERK-/- and MEFGCN2-/- cells.

In the last part of this report, we tested the chemotherapeutic agent sensitivity of

these four cell lines in hypoxia. Our results demonstrated that PERK and GCN2

contribute to the resistance to the treatment, but mutation of eIF2α did not impair the

effect.

Together, our findings are important for basic research and clinical research. It is

significant to find out if any impaired phosphorylation of eIF2α on serine 51 can hamper

hypoxia induced cell cycle arrest. The disruption of PERK or GCN2 can promote

hypoxia and chemotherapy sensitivity through a novel mechanism. The future

investigation will be focused on the effects of knocked down PERK or GCN2 on tumor

growth. In future studies, RNAi could be utilized to knock down the level of PERK and

GCN2 in cancer cells and cell viability will be investigated in hypoxia. In vivo studies can also be performed to investigate the roles of PERK and GCN2 in hypoxia. 89

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APPENDIX: LIST OF ABBREVIATIONS

1. ARNT: Aryl hydrocarbon nuclear receptor translocator

2. bHLH: Basic helix-loop-helix motif

3. CHX: Cycloheximide

4. C-terminus: Carboxyl terminus

5. CXCR4: CXC chemokine receptor 4

6. DAPI: 4',6-Diamidino-2-phenylindole

7. DMEM: Dulbecco’s modified eagle’s medium

8. DNA: Deoxyribonucleic acid

9. eIF: Eukaryotic initiation factors

10. EPO: Erythropoietin

11. ETS1: V-ets erythroblastosis virus E26 oncogene homolog 1

12. GCN2: General control non-depressible protein kinase 2, GCN2

13. GEF: Guanine nucleotide exchange factor

14. GLUT: Glucose transporter

15. GDP: Guanosine diphosphate

16. GTP: Guanosine-5'-triphosphate

17. HDM2: Human double minute 2 protein

18. HIF-1: Hypoxia inducible factor – 1

19. HIF-1α: Hypoxia inducible factor – 1 alpha

20. HK-1: Hexokinase 1

21. HRE: Hypoxic responsive element 109

22. HRI: Heme-regulated inhibitor kinase

23. LDH: Lactate dehydrogenase

24. Mdm2: Murine double minute 2 protein

25. NLS: Nuclear localization sequence

26. N-terminus: Amino terminus

27. ODD: Oxygen-dependent-degradation domain

28. PAS: Per-ARNT-Sim

29. PBS: Phosphate buffered saline

30. PBS-T: Phosphate buffered saline with Tween 20

31. PERK: RNA dependent protein kinase-like ER kinase

32. PKR: dsRNA induced protein kinase

33. pVHL: von-Hippel_Lindau protein

34. RNA: Ribonucleic acid

35. SDF-1: Stromal cell–derived factor-1

36. TAD: Transactivation domains

37. TGF-β: Transforming growth factor beta

38. UV: Ultraviolet

39. VEGF: Vascular endothelial growth factor